Synaptic PRG-1 Modulates
Excitatory Transmission
via Lipid Phosphate-Mediated Signaling
Thorsten Trimbuch,1,10 Prateep Beed,2,10 Johannes Vogt,1,10 Sebastian Schuchmann,2 Nikolaus Maier,2
Michael Kintscher,1,2 Jörg Breustedt,2 Markus Schuelke,3 Nora Streu,1 Olga Kieselmann,1 Irene Brunk,4 Gregor Laube,4
Ulf Strauss,1 Arne Battefeld,1 Hagen Wende,5 Carmen Birchmeier,5 Stefan Wiese,6 Michael Sendtner,6 Hiroshi Kawabe,7
Mika Kishimoto-Suga,7 Nils Brose,7 Jan Baumgart,1 Beate Geist,1 Junken Aoki,8 Nic E. Savaskan,1 Anja U. Bräuer,1
Jerold Chun,9 Olaf Ninnemann,1 Dietmar Schmitz,2,11 and Robert Nitsch1,11,*
1Institute
of Cell Biology and Neurobiology and NeuroCure
Research Center and NeuroCure
3Department of Neuropediatrics and NeuroCure Clinical Research Center
4Institute of Integrative Neuroanatomy
Charité, Universitätsmedizin Berlin, Charitéplatz 1, 10117 Berlin, Germany
5Max-Delbrück-Center for Molecular Medicine, Robert-Rössle-Str. 10, 13092 Berlin, Germany
6Institute of Clinical Neurobiology, Universität Würzburg, Zinklesweg 10, 97078 Würzburg, Germany
7Department of Molecular Neurobiology, Max Planck Institute of Experimental Medicine, Hermann-Rein-Str. 3 37075 Göttingen, Germany
8Graduate School of Pharmaceutical Sciences, Tohoku University, 6-3, Aoba, Aramaki, Aoba-ku, Sendai, Miyagi, 980-8578, Japan
9The Scripps Research Institute, 10550 North Torrey Pines Rd., La Jolla, CA 92037, USA
10These authors contributed equally to this work
11These authors contributed equally to this work
*Correspondence: robert.nitsch@charite.de
DOI 10.1016/j.cell.2009.06.050
2Neuroscience
SUMMARY
Plasticity related gene-1 (PRG-1) is a brain-specific
membrane protein related to lipid phosphate phosphatases, which acts in the hippocampus specifically at the excitatory synapse terminating on glutamatergic neurons. Deletion of prg-1 in mice leads
to epileptic seizures and augmentation of EPSCs,
but not IPSCs. In utero electroporation of PRG-1
into deficient animals revealed that PRG-1 modulates excitation at the synaptic junction. Mutation of
the extracellular domain of PRG-1 crucial for its interaction with lysophosphatidic acid (LPA) abolished
the ability to prevent hyperexcitability. As LPA application in vitro induced hyperexcitability in wild-type
but not in LPA2 receptor-deficient animals, and
uptake of phospholipids is reduced in PRG-1-deficient neurons, we assessed PRG-1/LPA2 receptordeficient animals, and found that the pathophysiology observed in the PRG-1-deficient mice was
fully reverted. Thus, we propose PRG-1 as an important player in the modulatory control of hippocampal
excitability dependent on presynaptic LPA2 receptor
signaling.
INTRODUCTION
Recently, we discovered a new class of proteins, which we
named plasticity-related genes (PRGs) because two members
1222 Cell 138, 1222–1235, September 18, 2009 ª2009 Elsevier Inc.
of the PRG family, PRG-1 (LPR-4) and PRG-3 (LPR-1), play
a role in regulating cellular plasticity. Their actions include filopodia formation, neurite extension, axonal path finding and reorganization after lesions (Bräuer et al., 2003; Peeva et al., 2006;
Savaskan et al., 2004; Sigal et al., 2007). PRGs are integral
membrane proteins that possess homology to lipid phosphate
phosphatases (LPPs) and can be classified as part of the superfamily of lipid phosphatases/phosphotransferases (PAP2, also
known as LPT) (Sigal et al., 2005). PRG-1 bears the characteristic
LPP feature of three conserved domains facing the extracellular
side of the plasma membrane or the luminal side of cellular
membranes (Zhang et al., 2000). These domains enable the
LPPs to dephosphorylate a variety of phosphorylated lipid
substrates, such as phosphatidate (PA), lysophosphatidate
(LPA) or sphingosine-1-phosphate (S1P) (Brindley and Waggoner, 1998). Bioactive lipid phosphates are key factors in initiating
receptor-directed signaling cascades (Anliker and Chun, 2004;
Fukushima and Chun, 2001; Milstien et al., 2007) and participate
in diverse cellular processes, such as cell proliferation, neurite
retraction, cortical development, and regulation of exocytosis
(Kingsbury et al., 2003; Moolenaar et al., 2004; Pan et al., 2007;
Spiegel et al., 2002; Ye et al., 2002). Therefore, LPPs may act
as negative regulators of these signaling cascades dependent
on critical amino acids within the conserved domains (Brindley,
2004; McDermott et al., 2004; Neuwald, 1997; Pyne et al.,
2005; Sigal et al., 2005). Interestingly, unlike other members of
the PAP2 superfamily, PRG-1 contains an additional C-terminal
hydrophilic domain of around 400 amino acids (Bräuer et al.,
2003). We previously showed that the neuronal cell line
N1E-115, when transfected with a prg-1-GFP fusion construct
and exposed to LPA, generates increased extracellular LPA
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Figure 1. PRG-1 Is a Neuronal, Synaptic Molecule Expressed by Glutamatergic Neurons
(A) PRG-1 expression in the adult hippocampus as shown by immunohistochemistry (the scale bar represents 250 mm on the left, and 50 mm on the right, respectively). Dendrites of CA1 pyramids exhibit high expression, with membrane-bound expression shown in higher magnification (insets, arrows). PRG-1 did not
colocalize with glial markers such as GFAP (the scale bar represents 15 mm).
(B) In situ hybridization of prg-1 colocalized with the neuronal marker NeuN but not with GFAP+ cells in the CA1 region (the scale bar represents 50 mm).
(C) PRG-1/prg-1 was not expressed in interneurons as shown for GAD67, Parvalbumin (PV, arrows), Calbindin (CB, arrows) and Calretinin (CR) by immune stainings and in situ hybridization but was expressed in CB+ CA1 pyramids (arrowheads) (the scale bar represents 15 mm).
(D) PRG-1 was mainly expressed on dendrites and colocalized with ProSAP2 and Homer, but not Gephyrin (the scale bar represents 2 mm). Subcellular fractionation of mouse cerebral cortex revealed PRG-1 within the PSD. RabGDI was slightly enriched in the SC+CSV fraction but undetectable in the PSD-95 enriched
PSD fraction, whereas PRG-1 was detected in this fraction but not in the presynaptic vesicle fraction (SC+CSV). Ultrastructural analysis of PRG-1 expression
shows preferential expression of PRG-1 at the postsynaptic density (the scale bar represents 200 nm).
degradation products. Moreover, these cells were protected
against LPA-induced neurite collapse in a dose-dependent
fashion (Bräuer et al., 2003; McDermott et al., 2004), which
demonstrates that PRG-1 had interfered with lipid phosphate
signaling. Our studies have so far revealed that expression of
prg-1 is vertebrate- and brain-specific, and that the gene is highly
conserved in vertebrate species, including humans. Expression
of PRG-1 begins perinatally, is strongly expressed in hippocampal neurons and rapidly upregulated after lesion of the glutamatergic entorhinal fiber tract (Bräuer et al., 2003).
RESULTS
Neuronal Localization of PRG-1
Using an antibody against PRG-1, we found the protein exclusively in neurons, as shown by colocalization with the neuronal
marker NeuN (Figure 1A). GFAP+ astrocytes or microglial cells
and oligodendrocytes (data not shown) were not stained
(Figure 1A). Double staining and in situ hybridization showed
expression of the prg-1 gene exclusively in NeuN-containing
neurons, but not in astrocytes (Figure 1B). PRG-1 did not
Cell 138, 1222–1235, September 18, 2009 ª2009 Elsevier Inc. 1223
colocalize with GABAergic markers including glutamic acid
decarboxylase-67 (GAD67) or calcium-binding proteins, such
as calbindin (CB), calretinin (CR) and parvalbumin (PV). Similarly,
no colocalization was observed with prg-1 mRNA by in situ
hybridization (Figure 1C). PRG-1 colocalized with ProSAP2 and
Homer1, molecules specifically located at the postsynaptic
density of glutamatergic synapses (Brakeman et al., 1997; Tu
et al., 1999; Wouterlood et al., 2003), but not Gephyrin, a marker
at the postsynaptic side of GABAergic contacts (Sassoe-Pognetto and Fritschy, 2000). Subfractionation of the synaptosomal
preparation (Cotman and Taylor, 1972) showed that PRG-1 was
present in the postsynaptic density (PSD), but absent in presynaptic vesicles. Finally, localization at the ultrastructural level
revealed DAB-reaction product, indicating PRG-1 immunolabeling exclusively at postsynaptic structures of asymmetric synapses of CA1 pyramidal cells and silver-enhanced immunogold
signal of PRG-1 bordering the synaptic cleft (Figure 1D). These
data indicate that PRG-1 plays a role at the postsynaptic site
of glutamatergic neurons in the hippocampus.
EEG Recordings Reveal Seizure Activity
in PRG-1 KO Mice
We disrupted the prg-1 gene by inserting an IRES-LacZ-MC1Neo cassette, which deletes the extracellular domains coded
on exons 5 and 6 and blocks translation of the intracellular tail
coded by exon 7 (Figure S1 available with this article online).
The PRG-1 / mice were viable but showed a transiently
reduced weight during development (Figure S2A), a reduction
in brain weight (Figure S2B), and a mortality of approximately
50% at 3-4 weeks after birth (Figure S2C). Since we observed
seizure-like movements of some PRG-1 / mice around postnatal day (P) 20, we made in vivo electrographic recordings using
intracranial, epidural electrodes above the somatosensory
cortex in knockout (KO) and wild-type (WT) mice at P19-P31.
In all investigated KO mice (n = 10), but none of the WT mice
(n = 7), hypersynchronized activity and preictal events were identified by field potential shifts (>150 mV) as early as around P20
(Figure 2A). Electrographic ictal events and tonic-clonic seizures
were observed in 5 of 10 KO mice over the following two days
(Figures 2A–2C and Movies S1 and S2). The frequency of field
potential discharges was 3–5 Hz during ictal activity and
0.8–1.6 Hz during interictal activity (Figure 2B). All recorded KO
mice exhibiting electrographic seizures died during status
epilepticus (ictal activity > 30 min). The KO mice with implanted
electrodes at both hemispheres showed preictal events at only
one, but ictal activity at both, hemispheres (KO, n = 4; WT, n = 3;
Figure 2C), suggesting a focal ictogenesis with secondary generalization. In a number of KO mice only hypersynchronized
activity, but no ictal events, were registered (n = 3). These in vivo
recordings indicate that lack of PRG-1 results in severe alteration
of synaptic transmission leading to juvenile epileptic seizures.
No Evident Structural Changes, but Pathological
Network Properties in Hippocampal Slices
Surprisingly, histological and immunocytochemical studies
revealed no morphological alterations in PRG-1 / mice. In the
hippocampus of PRG-1 / mice at P21, pyramidal neurons
had developed normally as shown by the neuronal marker
1224 Cell 138, 1222–1235, September 18, 2009 ª2009 Elsevier Inc.
NeuN (Figure 3A). Specifically, the distribution of inhibitory
GABAergic interneurons expressing parvalbumin was not
altered in these mice (Figure 3A). Also, synaptic innervation of
CA3 and CA1 pyramidal cells was unchanged at P21 in
PRG-1 / mice (Figure S3). Whole-cell patch clamp experiments
in current clamp mode on WT and KO mice at that age did not
reveal any obvious difference in intrinsic properties of CA1 pyramidal cells that would indicate alteration of the cellular architecture of individual neurons (Figure 3B). In vitro studies of the
robust network rhythms in the hippocampus (Maier et al.,
2003) by analyzing sharp waves (SPW) and accompanying
ripples in the CA3 area in hippocampal slices from PRG-1 KO
mice at P21 (Figure 3C) revealed that SPW incidence was not
altered in either condition (WT: 2.2 ± 0.2 Hz versus KO: 2.4 ±
0.6 Hz; n = 8 for both groups, Mann-Whitney Test, p = 0.44;
Figure 3D). Likewise, spectral frequency and ripple power were
not significantly altered in slices from WT or PRG-1 KO animals
(frequency: 222.5 ± 4.5 Hz and 218.2 ± 6.8 Hz, for WT and KO,
respectively; n = 8, Mann-Whitney Test, p = 0.64; ripple power:
controls: 150 ± 43 mV versus 209 ± 48 mV, n = 8, Mann-Whitney
Test, p = 0.44; Figure 3D). However, bath application of carbachol induced gamma oscillation in WT slices (Figure 3E) (Fisahn
et al., 1998). In slices from KO animals, we additionally detected
interictal discharges (78% of tested slices; compare to data
presented in Figure 3C with summary diagram in Figure 3D).
These findings suggest that PRG-1 deficiency results in pathological network synchronization, probably due to a pathological
increase of excitatory synaptic transmission.
Hyperexcitability in CA1 Neurons
To elucidate the mechanistic role of PRG-1 in the cellular physiology of hippocampal pyramidal neurons, we first analyzed the
so-called input-output behavior in area CA1 of acute hippocampal slices around P21. fEPSPs evoked by Schaffer collateral
stimulation in the stratum radiatum of the CA1 hippocampal
region showed that slices from PRG-1 KO mice (n = 10) were
highly hyperexcitable compared with WT litters (n = 3, paired
t test for every fiber volley, p < 0.05) (Figure 4A). PRG-1+/ mice
(n = 6), which show a PRG-1 expression of approximately 50%
of that of WT animals, exhibited intermediate values (Figure 4A).
At single-cell level, PRG-1 KO mice showed a larger synaptic
current (Figure 4B) and significantly higher mEPSC frequency
(KO (n = 15, 3.22 ± 0.34 Hz) compared to WT mice (n = 14,
1.79 ± 0.26 Hz); student’s t test: p < 0.01; Figure 4C, left panel)
as recorded from CA1 pyramidal cells in voltage clamp mode.
However, the mEPSC amplitude did not vary significantly
between PRG-1 KO and WT (Kolmogorov-Smirnov test, p > 0.05,
WT mean = 16.63 ± 0.13 pA, KO mean = 15.6 ± 0.14 pA,
Figure 4C, right panel). This effect was specific for excitatory
transmission on pyramidal neurons because (1) synaptically
evoked IPSCs at different stimulation strengths did not differ
between WT (n = 8) and PRG-1 KO (n = 7; paired t test for every
stimulation strength, not significant for all points with p > 0.05;
Figure 4D) mice; (2) the IV-curve of inhibitory currents showed
no significant difference between WT (n = 6) and PRG-1 KO
(n = 5, Figure 4E) mice, and (3) mIPSC frequency (WT n = 4,
2.13 ± 0.08 Hz, KO n = 4, 1.96 ± 0.07 Hz; student’s t test:
p = 0.13; Figure 4F) and amplitude (Kolmogorov-Smirnov test,
Figure 2. PRG-1 KO Mice Show Epileptic Activity at the End of the Third Week of Life
(A) Preictal events at P20/P21 and ictal activity at P22 during in vivo recordings of a PRG-1 KO mouse (left column; right column: WT littermate). Video images
show tonic-clonic seizures of the same KO mice at P22.
(B) Ictal (red) and interictal (blue) periods during in vivo electrographic recording in a P21 PRG-1 KO mouse.
(C) In vivo recording of a PRG-1 KO mouse with implanted electrodes at both hemispheres at P20-P22. Hypersynchronized activity and preictal events can be
identified at the right hemisphere around P20/P21 (see arrows); generalized ictal activity is seen over both hemispheres at P22.
p > 0.05, WT mean = 13.85 ± 0.07 pA, KO mean = 13.01 ± 0.1 pA,
Figure 4F, right panel) did not vary between WT and PRG-1 KO
mice. These findings demonstrate that excitatory transmission
in PRG-1 KO mice was significantly increased in CA1 pyramidal
neurons, whereas inhibitory events were not altered.
PRG-1 Action in Single CA1 Neurons
To differentiate between network and single-cell phenotype, the
prg-1 gene was deleted in a small subset of CA1 pyramidal cells
that normally express PRG-1 protein (Figure 1A). This was
achieved by Cre recombinase expression, which coexpressed
GFP in a PRG-1-conditional KO mouse (for construct, see
Figure S4) using in utero electroporation. These animals, aged
about P21, did not show any signs of seizures. We performed
simultaneous recordings from GFP+ (PRG-1 KO genotype) and
GFP (WT genotype) CA1 pyramidal neurons in acute hippocampal slices. Stimulus-evoked EPSC amplitudes were significantly increased in KO pyramidal cells compared with WT
Cell 138, 1222–1235, September 18, 2009 ª2009 Elsevier Inc. 1225
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Figure 3. No Evident Structural Changes, but Pathological Network Properties in Hippocampal Slices
(A) Juvenile PRG-1 KO mice (P21) show no changes in morphology as shown by NeuN staining or PV+ interneuron distribution in the hippocampus. The scale bar
represents 100 mm (left) and 50 mm (right).
(B) Resting membrane potential, input resistance and action potential height of CA1 pyramidal cells did not vary significantly between WT and PRG-1 KO mice.
(C) Hippocampal sharp wave (SPW)-ripples were expressed in PRG-1 KO slices. Upper traces: example data from CA3 in a WT (black trace) and KO slice (red
trace). Below: magnified signals (see asterisks for respective locations in recordings), their band-pass-filtered derivative and SPW peak-triggered averages
(±SEM), respectively, of 50 randomly selected SPWs.
(D1) SPW-incidence was not affected in either condition.
1226 Cell 138, 1222–1235, September 18, 2009 ª2009 Elsevier Inc.
pyramidal cells (KO (n = 7); WT (n = 7); student’s t test: p < 0.01;
Figure 5A, left). Recordings from GFP+ and GFP CA1 pyramidal
cells showed a significantly higher mEPSC frequency in the
PRG-1-deficient pyramidal cells compared with PRG-1-expressing pyramidal cells (KO (n = 7, 4.02 ± 0.78 Hz); WT (n = 4, 1.96 ±
0.23 Hz); student’s t test: p < 0.05; Figure 5B left). In this setting,
the majority of cells expressed PRG-1, as did, consequently, the
majority of afferents on a PRG-1 / CA1 pyramidal cell derived
from WT CA3 neurons. This indicates that the increase in neuronal
excitability is due to the lack of PRG-1 at the postsynaptic side.
Next, we electroporated prg-1 coexpressing GFP+ into
PRG-1 KO mice to analyze whether expression of PRG-1 in
single neurons can rescue the pathophysiological phenotype
independently of the surrounding KO environment. Simultaneous recordings from GFP+ (PRG-1+) and GFP (KO genotype)
CA1 pyramidal neurons in acute hippocampal slices showed
significantly decreased EPSC amplitudes in PRG-1-expressing
pyramidal cells compared with KO pyramidal cells (PRG-1+/
GFP+ (n = 10); KO/GFP (n = 10); student’s t test: p < 0.01;
Figure 5A right). mEPSC frequency was significantly reduced in
the GFP+ cells expressing PRG-1, indicating a rescued function
(KO/GFP : n = 4, 4.01 ± 0.29 Hz; PRG-1+/GFP+: n = 5, 1.87 ±
0.08 Hz; student’s t test: p < 0.01; Figure 5B right). These data
show that lack of PRG-1 in an individual postsynaptic neuron
increases its excitatory synaptic input.
No Evidence for Changes in the Postsynaptic Molecular
Machinery at the Synapse
To test for changes in the molecular machinery of synaptic transmission, we analyzed presynaptic markers, such as the vesicular
GABA or glutamate transporters and postsynaptic AMPA and
NMDA receptors subunits. Immunofluorescent staining showed
no alteration in the expression of these molecules between PRG1 KO and their WT litters within the hippocampus at P21
(Figure 6A). Western blotting showed no changes in expression
of the postsynaptic density marker PSD-95 or the glutamate
receptor subunits NR-1, NR2A/B, GluR1, GluR2/3, and GluR4
at the postsynaptic density in PRG-1 KO mice compared with
WT littermates (Figure 6B). We used an antibody against an
extracellular epitope of GluR2 as a marker for AMPA receptors
located in the dendritic membrane, which were in direct colocalization with VGlut1 as a marker for glutamatergic synapses. We
could not detect any significant difference in the presence of
AMPA receptors at glutamatergic synapses (Figure 6C). Finally,
we studied the AMPA/NMDA ratio (Figure 6D) and changes in
the holding current in postsynaptic neurons on AMPA application (Figure 6E), both of which showed no difference between
WT and PRG-1 KO animals (AMPA/NMDA: n (WT) = 4, n (KO) = 8,
p = 0.75; AMPA wash in: n (WT) = 4, n (KO) = 5, p = 0.53).
Therefore, we concluded that lack of PRG-1 was not affecting
the postsynaptic molecular machinery involved in excitatory
transmission, which would have resulted in overexcitation and
eventually seizure activity. Rather, we hypothesized a postpresynaptic signaling pathway that is modulated by PRG-1.
PRG-1 Interacts with Bioactive Phospholipids
Next, we assessed the potential interaction of PRG-1 with bioactive phospholipids such as LPA at the synapse. Using fluorescence-coupled phosphatidic acid, as established in expression
systems by other groups (Renault et al., 2004; Roberts and
Morris, 2000), uptake of fluorescence-labeled lipids was reduced
by about 50% in PRG-1 / neurons when compared to WT
neurons in culture (n (WT) = 30, n (KO) = 38, student’s t test,
p < 0.001) (Figure 7A). We then determined whether PRG-1
modulation of excitability at glutamatergic synapses is dependent on its involvement in lipid phosphate signaling by expressing
a mutated PRG-1 protein in PRG-1 / animals using in utero
electroporation. This protein was coexpressed with GFP+, and
point-mutated at a critical extracellular motif (H253K) known to
disrupt the interaction of LPPs with lipid phosphates in the
extracellular space (Bräuer et al., 2003; McDermott et al., 2004;
Starz-Gaiano et al., 2001; Zhang et al., 2000). Although the
protein was also detected at the plasma membrane of GFP+ pyramidal cells at glutamatergic synapses (Figure S6), mEPSC
frequency was not reduced, indicating that a functional rescue,
as obtained by expression of the WT protein, was not achieved
(GFP+ (n = 8, 3.04 ± 0.29 Hz); GFP (n = 4, 2.96 ± 0.23 Hz);
student’s t test: p = 0.855; Figure 7B). Taken together, these findings suggest that the modulatory role of PRG-1 in excitatory
transmission depends on its interaction with and uptake of bioactive lipids mediated by the molecule’s extracellular domains.
PRG-1 Acts via LPA2 Receptors at the Excitatory
Synapse
Finally, we investigated the signaling mechanisms at the presynaptic terminal modulated by PRG-1 at the excitatory synapse.
In an independent set of experiments, we directly tracked the
time course of the LPA influence on mEPSCs on single CA1 pyramidal cells in slices of WT animals. Here, bath application of
10 mM LPA induced a two-fold increase in mEPSC frequency
in each CA1 neuron of WT animals tested (WT: before LPA,
0.98 ± 0.20 Hz; after LPA, 1.86 ± 0.32 Hz; mean ratio LPA/baseline: 2.00 ± 0.11; n = 10; p < 0.01; paired student’s t test;
Figure 7C). In contrast, no change was detected in mIPSC
frequency (Figure S7A). Moreover, in PRG-1 / animals, LPA
enhanced the mEPSC frequency to a mean ratio of 1.34 ± 0.08
(n = 7, p < 0.01). This increase, however, was significantly smaller
in comparison to WT animals (p < 0.001, independent t test;
Figure S7B).
LPA did not induce these effects in slices obtained from
LPA2 receptor (LPA2-R)-deficient animals (LPA2-R-KO: before
LPA 0.67 ± 0.20 Hz, after LPA 0.64 ± 0.21 Hz; mean ratio
(D2) Averaged PSD functions (±SEM) reveal unchanged spectral frequency of ripple oscillation in WT and KO slices. Right: Unaltered ripple power in PRG-1 / .
(D3) Distribution of slices expressing different classes of hippocampal network oscillations: all tested slices expressed SPWs (100%); following carbachol administration, 75% of slices from WT and 62.5% of slices from PRG-1 / animals displayed gamma activity. Interictal-like discharges were not observed in WT but
frequently in KO slices (77.8%). Numbers indicate slice fractions displaying the respective electrographic events.
(E) Example traces showing gamma oscillation from a WT slice (left) and seizure-like events in a KO slice (right) following carbachol administration, with magnified
example (box).
Cell 138, 1222–1235, September 18, 2009 ª2009 Elsevier Inc. 1227
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Figure 4. Hyperexcitability in CA1 Neurons in PRG-1 KO Mice
(A) Extracellular fEPSPs were evoked by Schaffer collateral stimulation. fEPSP slope is plotted for WT, PRG-1+/ and PRG-1 / . Increased network excitability is
seen for the PRG-1 / mice (white) compared with WT mice (black). In WT, PRG-1+/ and PRG-1 / mice, the fEPSP slope increased linearly with increasing fiber
volley amplitudes, indicating a gene-dose-dependent effect of PRG-1 reflected in the adjacent western blot of brain homogenates.
(B) Evoked synaptic glutamatergic currents (at similar stimulation intensities and stimulation electrode positions) were enhanced for PRG-1 KO mice compared
with WT mice.
(C) A significantly higher mEPSC frequency was observed in PRG-1 KO CA1 pyramidal cells compared to WT cells (left panel), however the amplitude of these
events did not significantly vary between the two groups (right panel).
(D) Synaptically evoked IPSCs at different stimulation strengths did not differ between the WT and PRG-1 KO cells.
1228 Cell 138, 1222–1235, September 18, 2009 ª2009 Elsevier Inc.
EPSC in Cre – IUE
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mEPSC
15 pA
mEPSC frequency [Hz]
4
PRG-1
3
2
1
0
WT
+ Cre
GFP
15 pA
1s
merge
mEPSC frequency [Hz]
*
+ PRG-1
+ PRG-1
GFP
5
**
0.5
+ Cre
mEPSC
+ Cre
1.5
5
1s
4
PRG-1
3
**
2
1
0
KO
+ PRG-1
merge
Figure 5. PRG-1 Action in Single CA1 Neurons in Acute Slices
Using in utero electroporation prg-1 was deleted (GFP+) in conditional or re-expressed (GFP+) in constitutive PRG-1 KO mice in a subset of cells.
(A) Left: In simultaneous recordings (upper image) from GFP+ (KO) and GFP (WT, arrow) CA1 pyramidal neurons (middle image) EPSC amplitude (experimental
setting: lower image) was significantly increased in KO cells compared to WT cells. Right: In simultaneous recordings from GFP+ (PRG-1+) and GFP (KO) CA1
pyramidal neurons EPSC amplitude was significantly decreased in PRG-1-expressing cells compared to KO cells.
(B) Left: mEPSCs were recorded from GFP+ (KO) and GFP (WT) cells. Significantly higher mEPSC frequency was observed in the PRG-1 / cells compared to
WT neurons. Arrow points to GFP+ cell, in which PRG-1 (red) has been deleted. Right: mEPSC frequency was significantly reduced in the GFP+ (PRG-1-expressing KO) cells compared to the GFP (KO) cells. GFP signal in neurons indicates PRG-1 expression, confirmed by PRG-1 (red) immunolabeling.
Data are represented as mean ± SEM *p < 0.05, **p < 0.01. The scale bar represents 15 mm.
LPA/baseline: 0.96 ± 0.09; n = 6; p = 0.43; Figure 7C). We also assessed the presence of the LPA2-R in the hippocampus using
a specific antibody (Shida et al., 2004; Figure S8A). By means
of confocal triple labeling (Figure 7D), we identified specific
expression of LPA2 receptors in presynaptic terminals of the glutamatergic synapse due to its colocalization with Munc 13-1,
a marker for the active zone at the presynaptic membrane
(Varoqueaux et al., 2002), and VGlut-1, a marker for vesicles of
the glutamatergic bouton (Takamori et al., 2000; Figures 7D
and S8B). Immunostaining for LPA2-R was found in presynaptic
terminals of an asymmetric, glutamatergic junction, while
symmetric, GABAergic synapses in the direct vicinity were not
labeled (Figure 7E). As final proof of our hypothesis, we studied
excitatory events in PRG-1/LPA2–R / mice. fEPSPs evoked by
(E) IV curves of inhibitory currents show no significant difference between the WT and PRG-1 KO cells.
(F) mIPSC frequency (left panel) and amplitude (right panel) did not differ between PRG-1 KO and WT CA1 pyramidal cells.
Data are represented as mean ± SEM, **p < 0.01.
Cell 138, 1222–1235, September 18, 2009 ª2009 Elsevier Inc. 1229
WT
A
VGlut1
VGat
GAD 67
GluR1
GluR2/3
GluR4
NR1
Syn
VGlut1
VGat
GAD 67
GluR1
GluR2/3
GluR4
NR1
KO
Syn
B
E
D
WT
PRG-1-KO
50 pA
50 mS
10
AMPA/NMDA ratio
Colocalization per 10 μm2
GluR2ext / VGluT1
2.0
1.5
1.0
0.5
8
6
4
2
WT
PRG-1-KO
VGluT1 / GluR2ext
50
WT
PRG-1-KO
0
-50
-100
-150
0
0.0
Holding current [pA]
C
WT
PRG-1-KO
Baseline
AMPA Washout
Figure 6. No Evidence for Changes in the Classical Molecular Machinery at the Synapse
(A) Immunofluorescence for presynaptic markers, such as synaptophysin (Syn), vesicular glutamate transporter 1 (VGlut1), vesicular GABA transporter (VGat) and
glutamate decarboxylase 67 kDa isoform (GAD67), as well as postsynaptic markers, such as AMPA receptor subunits (GluR1, Glur2/3, GluR4) and the NMDA
receptor subunit NR1, do not differ between PRG-1 KO and WT litters. The scale bar represents 50 mm.
(B) Protein analysis of the PSD fraction prepared from PRG-1 KO and WT mice. 2.5 mg of purified PSD fractions were subjected to western blotting to estimate
expression levels of NR1, NR2A/B, and GluR1, GluR2/3, GluR4, and PSD-95. Expression of the synaptic proteins were quantified and normalized by the loading
control (b-actin) and to the WT.
(C) Immune staining for VGluT1 and GluR2ext on dendrites of CA1 pyramidal cells in WT and PRG-1 KO mice. Arrows indicate colocalization. Stereological analysis of synaptically localized GluR2 revealed no difference between WT (n = 5) and PRG-1 KO (n = 5).
(D) The AMPA/NMDA ratio did not differ between the WT and PRG1-KO mice.
(E) The holding current changes on AMPA (50 nM) wash-in did not differ significantly between the WT and PRG-1 KO mice.
Data are represented as mean ± SEM.
1230 Cell 138, 1222–1235, September 18, 2009 ª2009 Elsevier Inc.
Schaffer collateral stimulation in the stratum radiatum of the CA1
hippocampal region from double-deficient mice (n = 7) were not
hyperexcitable in comparison to slices from WT litters (n = 3,
paired t test for every fiber volley, p > 0.05), which was in contrast
to findings in single PRG-1 / animals (Figures 7F and 4A). On
single-cell level, mEPSCs from CA1 pyramidal cells showed
a significantly reduced frequency in PRG-1/LPA2–R / mice
(n = 7, 1.60 ± 0.13 Hz) compared to that of PRG-1 / mice
(n = 8, 3.48 ± 0.22 Hz; student’s t test: p < 0.001; Figure 7F), which
was in the range of WT animals (see Figure 4C). These doubledeficient animals showed neither reduced body weight during
the postnatal phase nor seizure-like behavior or increased
mortality around P21 as seen in PRG-1 / mice (Figure S2A,
S9). Finally, in vivo EEG-recordings revealed that, unlike single
PRG-1 KO mice, double-deficient animals showed no signs of
epileptic discharge around P21 (Figure 7G and Movie S3;
n = 4). These experiments demonstrated that changes to hippocampal network excitability, increase in glutamatergic transmission and seizure activity are fully reverted in the absence of the
LPA2-R.
DISCUSSION
Our results show that deletion of the LPP-related molecule
PRG-1 results in severe hippocampal overexcitability and leads
to epileptic discharge in juvenile animals. This is due to a specific
role of PRG-1 at the excitatory synapse on glutamatergic
neurons, which appears to be dependent on the interaction
between PRG-1 and lipid phosphates such as LPA, able to signal
to presynaptic LPA2–receptors. These data indicate a scenario
for the regulation of glutamatergic transmission controlled by
PRG-1, which involves mechanisms mediated by bioactive lipids
such as LPA and supplements the classical molecular machinery
at the synapse.
PRG-1 expression on dendrites is limited to glutamatergic
neurons and reaches maximum levels in the third week of life
(data not shown), a time when synaptogenesis peaks and
neuronal connections are shaped. In parallel, expression of
LPA2–receptors is high (Contos and Chun, 2000; Contos et al.,
2000) at this critical period of seizure susceptibility/increased
hyperexcitability (Ben-Ari et al., 2007), in which an imbalance
between excitatory and inhibitory inputs in the almost developed
neurons is believed to occur (Ben-Ari, 2006). The lack of
apparent morphological changes in PRG-1 KO animals, and
specifically the presence of GABAergic neurons as in WT
animals, indicated that changes in network properties were
due to a pathological increase of excitatory synaptic transmission on otherwise unaffected neurons. Field potential recordings
of hippocampal slices confirmed this, showing increased excitability in PRG-1 KO mice while intrinsic properties of the CA1
neurons and the overall morphology were not altered.
The fact that PRG-1 / CA1 pyramidal cells showed a significantly higher stimulus-evoked EPSC amplitude and increased
mEPSC frequency while neither the stimulus-evoked IPSC
amplitude nor mIPSCs were affected by prg-1 deletion indicates
specific changes to glutamatergic transmission at the synapse.
However, future studies are required to determine whether glutamatergic synapses outside the hippocampus are also altered.
Our rescue of normal excitatory events in single PRG-1 / cells
by PRG-1 in utero electroporation demonstrates PRG-1’s effect
on neuronal excitability at individual glutamatergic synapses. In
turn, deletion of prg-1 in single cells of conditional PRG-1 KO
mice using Cre-recombinase in utero electroporation was sufficient to elicit the synaptic dysfunction. Thus, our data reveal
that hyperexcitability in PRG-1 KO animals can only be explained
by the lack of the modulatory activity of PRG-1 at the synapse.
As this single-cell KO approach never resulted in epileptic
discharge in the animals and the recorded PRG-1 KO neurons
were connected by normal presynaptic afferents deriving from
PRG-1-expressing cells, these findings confirm that PRG-1
influences excitatory transmission from the postsynaptic side.
Additionally, AMPA receptors at glutamatergic synapses and
their physiological function appeared to be normal. Therefore,
increased excitation at glutamatergic synapse, so far studied
in CA1 and CA3 (data not shown) in vitro, and related hyperexcitability and seizures in vivo, both caused by lack of postsynaptic
PRG-1, could not be explained by alteration in synaptic innervation or changes to postsynaptic receptors.
Thus, we addressed a direct role of PRG-1 in terms of its relation to lipid phosphate phosphatases. LPPs have been proposed
as negative regulators of receptor-directed signaling cascades
induced by bioactive lipid phosphates such as LPA or S1P
(Brindley, 2004; Sigal et al., 2005). However, little is known about
the physiological role of these molecules in the living organism,
particularly the brain. A function for germ cell migration of one
class of LPP homologs in vivo has been inferred from deletion
and ectopic expression experiments in Drosophila (Renault
et al., 2004; Starz-Gaiano et al., 2001). Homology of the
membrane-spanning N-terminal part of PRG-1 to the LPP family
clearly indicates their evolutionary relationship and implies that
the binding partner or ligand of PRG-1 is a bioactive lipid such
as LPA (Todd et al., 2002). PRG-1 and LPPs share amino acids
shown to be crucial for the interaction of those molecules with
lipid phosphates. In particular, mutation of a highly conserved
histidine within the extracellular domain is known to abolish
this interaction (Starz-Gaiano et al., 2001; Zhang et al., 2000).
Precisely this exchange from histidine to lysine, when performed
in PRG-1 at position 253, resulted in a protein no longer able to
rescue the KO phenotype after in utero electroporation. These
data support the hypothesis that PRG-1 can interfere with glutamatergic signal transduction by interacting with lipid phosphates
in the synaptic cleft from the postsynaptic side (Figure S10).
Interestingly, the observed hyperexcitability in PRG-1 KO mice
was gene dose-dependent with heterozygous mice showing
input-output curves halfway between PRG-1-deficient and WT
litters. This approximately linear relationship between protein
expression and its electrophysiological action argues against
an enzymatic function of PRG-1. Moreover, PRG-1 contains
several nonconservative substitutions in the extracellular
domains involved in catalysis in other LPP family members,
which should hamper a relevant enzymatic activity of PRG-1
in vivo (McDermott et al., 2004). In the present study, we
detected that lack of PRG-1 in neurons significantly limits uptake
of bioactive lipids. Therefore, there is reason to assume that
apart from its potential residual enzymatic activity (Bräuer
et al., 2003), PRG-1 is effective in controlling the levels of LPA
Cell 138, 1222–1235, September 18, 2009 ª2009 Elsevier Inc. 1231
B
WT
C
PRG-1-KO
LPA2-KO
PRG-1 H253K
WT
+ LPA
PRG-1-KO
15 pA
0.8
***
0.6
0.4
0.2
0.0
WT
1s
3
LPA
mEPSC
frequency ratio
mEPSC frequency [Hz]
Relative fluorescent
intensity
4
1.0
2
1
2
1
0
-4
0
PRG-1-KO
20 pA
1s
+ LPA
KO
-2
H253K
0
2
4
6
mEPSC frequency ratio [LPA / control]
A
8
Time [min]
**
2.0
1,5
1,0
0,5
0
WT LPA2
KO
E
D
LPA2
VGlut1
Munc
F
fEPSP slope [mV / ms]
1.2
1.0
0.8
0.6
G
PRG-1-KO
WT
P20
PRG-1-KO
PRG-1-KO /
LPA2R-KO
P21
0.4
0.5 mV
0.2
50 ms
P22
0.0
0.00 0.05 0.10 0.15 0.20 0.25 0.30
Fiber volley [mV]
PRG-1-KO
PRG-1-KO / LPA2-KO
mEPSC frequency [Hz]
4
mEPSC
15 pA
1s
PRG-1-KO / LPA2-KO
3
2
P20
***
1
0
P21
P22
PRG-1 PRG-1-KO
KO
LPA2-KO
Figure 7. PRG-1 Influences Phospholipid Signaling
(A) NBD-PA uptake experiments in primary neurons revealed significantly lower fluorescence intensity in KO cells compared to WT litter cells, indicating reduced
lipid uptake (the scale bar represents 10 mM).
(B) Mutated PRG-1 (H253K) lacking the ability to interact with phospholipids was expressed in a subset of cells (GFP+) in PRG-1 KO animals using in utero electroporation. mEPSC frequency at P21 was not reduced to WT levels in PRG-1 (H253K)-expressing neurons after electroporation of the WT prg-1 construct (see
Figure 5).
(C) mEPSC frequency rapidly increased in CA1 pyramidal neurons after application of 10 mM LPA, but not in LPA2 receptor-deficient animals (LPA2-R-KO). For the
analysis, the mEPSC frequency of each analyzed neuron was averaged for the last four minutes under control conditions and in steady state after LPA treatment,
and these values then compared.
1232 Cell 138, 1222–1235, September 18, 2009 ª2009 Elsevier Inc.
at the synapse by nonenzymatic mechanisms. Thus, PRG-1
function may have evolved away from classical dephosphorylation performed by LPPs to mechanisms such as ‘‘sensor-like’’ or
receptor/transporter-like activities. This has been shown for
members of other enzyme families that have functions different
to those of enzymatic relatives due to mutations in critical catalytic residues in active sites (Todd et al., 2002).
The LPA2 receptor’s presynaptic location at the excitatory
junction and the fact that LPA does not increase excitability in
LPA2 receptor-deficient mice suggests this bioactive phospholipid is directly involved in controlling glutamatergic transmission. Consequently, the absence of both overexcitation at the
synapse and epileptic activity in PRG-1/LPA2 receptor-deficient
mice indicate PRG-1 modulates LPA-mediated control of excitatory discharge via the LPA2 receptor. Changes in lipid phosphate species have been linked to forms of human epilepsy
(Hermansson et al., 2005). Receptors for lipid phosphates have
been described (Fukushima and Chun, 2001) as acting via G
proteins and are putative transducers of a signal mediated by
bioactive lipids at the synapse. Furthermore, it has been
reported that lipid phosphates and their receptors can interfere
with excitability (MacLennan et al., 2001) and exocytosis via G
protein-coupled receptor-mediated signaling pathways (Pan
et al., 2007). This indicates that modulation of neuronal transmission might involve bioactive lipids such as LPA acting via presynaptic LPA2 receptors, a mechanism that is in turn controlled
by PRG-1 from the postsynaptic side specifically at the glutamatergic junction.
EXPERIMENTAL PROCEDURES
Generation of the Mouse Lines
Constitutive PRG-1 KO mice were generated by replacing exons 4 to 5 of prg-1
with an IRES-b-Gal-Neo cassette. Conditional PRG-1 KO mice were generated by flanking exons 4 to 6 of prg-1 with LoxP sites, allowing removal by
Cre recombinase expression. (see Supplemental Data). To generate a PRG1/LPA2 receptor / mouse line, heterozygous constitutive PRG-1 KO mice
were bred with the LPA2-R mouse line, provided by J. Chun (Contos et al.,
2002).
Immunohistochemistry and DIG In Situ Hybridization
Mice were anesthetized (0.5% ketamine i.p.) and transcardially perfused
with 4% PFA for IHC and DIG in situ. Brains were postfixed overnight in
4% PFA and further processed. Primary neurons or vibratome sections
were blocked for 1 hr (5% normal goat serum and 0.1% Triton X-100) before
overnight incubation with primary antibodies (see Supplemental Data).
Sections were immunostained with Alexa-labeled secondary antibodies (Invitrogen) or with biotinylated secondary antibodies (Vector Laboratories).
Ultrastructural analysis was performed as described (Krauss et al., 2007;
Lujan et al., 1997). DIG in situ hybridization was performed according to
manufacturer recommendations (Roche Applied Sciences) using a 400 bp
DNA fragment from exon 7.
Quantitative Analysis of GluR2
Analysis of synaptic GluR2 was performed using antibodies against an extracellular epitope (1:300 Chemicon [Rouach et al., 2005]) and against VGlut1
(1:1000; Synaptic Systems). Colocalization of GluR2 and VGlut1 along the
dendrites of CA1 neurons was analyzed by stereological methods. Briefly,
from each slice two optical sections (1 mm apart) were analyzed using an unbiased counting frame. Colocalized GluR2 and VGlut1 punctae were only
counted if also present in the lower optical section as described (Mokin and
Keifer, 2006).
Electrophysiology
We performed in vitro experiments, approved by the Berlin state government
(T0100/03), on slices of mice (21–28 days). Mice were briefly anesthetized with
isofluorane and decapitated. Brains were rapidly removed and transferred to
cooled (1 C–4 C) oxygenated artificial cerebrospinal fluid (ACSF), containing
(in mM): NaCl (87), NaHCO3 (26), sucrose (75), glucose (25), KCl (2.4), NaH2PO4
(1.25), MgCl2 (7), and CaCl2 (0.5); (pH 7.4). Horizontal hippocampal slices
(300 mm; 400 mm for oscillation experiments) were cut on a Leica VT 1200
Vibratome (Leica Microsystems, Germany). Slices were then incubated at
34 C–35 C for 30 min or transferred to an interface-type recording chamber
for oscillation experiments. All recordings were performed in ACSF containing
(in mM): NaCl (119), NaHCO3 (26), glucose (10), KCl (2.5), NaH2PO4 (1.25),
MgCl2 (1.3) and CaCl2 (2.5), at room temperature (31 C–33 C for interface
chamber experiments). All ACSF solutions were equilibrated with 95% O2
and 5% CO2; osmolarity of ACSF for recordings was maintained in the range
of 290 and 305 mosmol/l. Extracellular and whole-cell patch clamp recordings
in vitro were performed at 32 ± 1 C in either a submerged or interface-type
recording chamber. For these recordings, we used borosilicate glass electrodes (2-5 MU) filled with (mM): K-gluconate (135), HEPES (10), Mg-ATP (2),
KCl (20), EGTA (0.2); pH of the intracellular solution was adjusted to 7.2 with
KOH. For extracellular recordings, glass microelectrodes with a tip diameter
of 5 mm were filled with ACSF before use. In some interface experiments,
tungsten wire electrodes were used to sample hippocampal oscillations.
Principal cells located in the pyramidal cell layer were identified using infrared
differential interference contrast video microscopy. In the whole-cell configuration, de- and hyperpolarizing current steps (200 ms) were applied to characterize the cells’ intrinsic properties; only cells that showed typical spiking
characteristics of principal neurons were considered. Series resistance (Rs)
was monitored throughout experiments; cells were rejected if Rs was > 20 MU
or varied > ± 30% during the recording. No Rs compensation was used.
Extracellular signal was amplified 1000x and filtered at 1 Hz to 2 or 5 kHz.
All collected signals were digitized with 16-bit resolution (National Instruments,
Austin, USA) and sampled at 5 or 10 kHz using Igor Pro (Wavemetrics, Lake
Oswego, USA).
For electrographic recordings of cortical activity in vivo, teflon-coated silver
wire (uncoated diameter 75 mm; Goodfellow, UK) implanted at P17 or P18 was
used. The extracellular field signals were amplified 1000x (EXT 10-2F, npi,
Germany) and sampled at 3 kHz using a 16-bit data acquisition board PowerLab (8/30, ADInstruments, Spechbach, Germany) and the software Chart 5 Pro
(Version 5.5.6, ADInstruments). Data were offline high-pass filtered at 45 Hz
(for details see Supplemental Data).
(D) 3D reconstruction of a presynaptical terminal positive for the LPA2-R, the presynaptic glutamatergic marker VGlut1 and the active zone marker Munc13-1 (the
scale bar represents 0.5 mm).
(E) Ultrastructural analysis of the LPA2-R localization revealed a strong DAB signal on the presynaptic side of an asymmetric, presumably glutamatergic synapse
(arrowhead). Two adjacent axons display a positive signal underlining the presynaptic LPA2-R localization. In contrast, no LPA2-R signal was found in symmetric,
presumably inhibitory synapses (open arrowhead) (the scale bar represents 200 nm).
(F) Normal network excitability is seen in PRG-1 KO/LPA2-R KO / animals when compared to the PRG-1 KO mice. In contrast to PRG-1 KO mice, double KO
animals showed fEPSP slopes similar to WT animals. mEPSC frequency in CA1 pyramidal neurons in hippocampus from double KO mice also returned to WT
levels and did not show the increase seen in PRG-1 / animals.
(G) In vivo recordings of double KO animals did not display the hypersynchronized activity around P21 seen in PRG-1 / mice of same age.
Data are represented as mean ± SEM, **p < 0.01, ***p < 0.001.
Cell 138, 1222–1235, September 18, 2009 ª2009 Elsevier Inc. 1233
In Utero Electroporation
The in utero electroporation experiments were carried out as described (Prozorovski et al., 2008), in accordance with a protocol approved by the local
animal welfare committee. Briefly, we used time-pregnant mice at E15–E16
(post coitum). The uterine horns of anesthetized mice were exposed and the
DNA solution was injected through the uterine wall into the lateral ventricle
of two of the embryos. Five 38 V pulses of 50 ms were applied at 950 ms intervals by holding the injected brain with forcep-type electrodes connected to
a square–pulse generator (CUY 21 Edit, Unique Medical Imada, Miyagi,
Japan).
Phospholipid Uptake Experiment
Lipid uptake assays and visualization of fluorescent PA in fixed primary
neurons were performed using minor modifications of described methods
(Renault et al., 2004; Roberts and Morris, 2000). Primary hippocampal neurons
of WT and PRG-1 KO mice were prepared from embryonic day 18 mice as
described (Brandt et al., 2007). After 7 days in vitro neurons were washed in
serum-free culture medium and incubated with lipid vesicles composed of
20:5 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (PC) / 1-hexanoyl-2[6-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]hexanoyl]-sn-glycero-3-phosphate (NBD-PA) (10 mM total) (Avanti Polar Lipids, Alabaster, U.S.A.) for 5 min
at 37 C. Then the cells were placed on ice, washed once with PBS and fixed for
20 min with cold 4% paraformaldehyde and mounted in antifade reagent. Fluorescent neurons of WT and PRG-1 KO were scanned with a Leica confocal
laser-scanning microscope (SL) equipped with an Argon laser (excitation line
488 nm) and 40x oil objective (4x zoom). Same beam path configurations for
both WT and KO cells were employed. Fluorescent images were quantified
using ImageJ. Resulting mean values of fluorescent intensity were normalized
to WT.
Pharmacological Compounds
Carbachol, D-APV, TTX and GABAzine were added from aqueous stock
solutions; cyclothiazide and NBQX were administered from stock solutions
dissolved in DMSO (final concentration of DMSO < 0.1%). All drugs were
purchased from Tocris/BioTrends GmbH or Biozol GmbH (Enching, Germany)
and Sigma-Aldrich. A 2.6 mM LPA stock solution was prepared by dissolving
LPA (Sigma-Aldrich, Munich, Germany) in 50 mM HEPES, 138 mM NaCl,
2.7 mM KCl, 1 mM CaCl2, 1mM MgCl2 and 1% BSA and subsequently diluted
in ACSF.
Data and Statistical Analysis
Data analysis was performed using SigmaPlot (Systat Software Inc., San
Jose, USA), GraphPad Prism 4 (Graph Pad Software, La Jolla, USA) and
Matlab (The Mathworks, Aachen, Germany). Spontaneous miniature
EPSCs/IPSCs were detected using a threshold algorithm generated in
Matlab. For calculating AMPA/NMDA ratio, the NMDA current was measured
by holding the cell at +40 mV. 100 ms after the stimulus artifact, the amplitude
was read off as the NMDA component. SPW-R detection was based on
a voltage-threshold (>3.5SD of event-free baseline; see Supplemental
Data). PSD was computed in Matlab using Thomson’s multi-tapering method
with adequate window and overlap values (Thomson, 1982). To detect electrographic seizure activity in vivo we used a detection threshold to define the
time point when the ictal activity reached an amplitude > 3SD of activity
during baseline. Data are reported as mean ± SEM, which are also presented
as error bars in the figures. Statistical significance was assessed using
student’s t test, Kolmogorov-Smirnov test, Wilcoxon’s signed rank test or
the Mann-Whitney U-test for paired or unpaired data at the given significance
level (p).
ACKNOWLEDGMENTS
The authors thank James Ari Liebkowsky for correcting the manuscript; Denis
Lajkó, Katja Rösler, Berit Söhl-Kielczynski, and Bernd Hesse-Niessen for
technical assistance; and Franz Theuring for providing the IRES-LacZ-MC1Neo cassette. This work was supported by the DFG: SFB 665/B3 to R.N.,
SFB 665/Z to C.B., and grant MH51699 (National Institutes of Health) to J.C.
Bettina Holtmann, Elvira Rhode, Katja Becker, and Boris Jerchow are
acknowledged for their help with embryonic stem cell work and blastocyst
injection. T.T. and P.B. are fellows of the GRK 1123. S.W. is now at Molecular
Cell Biology Group, Ruhr-Universität Bochum, 44801 Bochum, Germany, but
work was performed in the lab of M.S. and was supported by the DFG: SFB
581/B1 and Z4 to M.S.
Received: July 31, 2008
Revised: April 30, 2009
Accepted: June 19, 2009
Published: September 17, 2009
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